Rapid casting of patterned vascular networks for
perfusable engineered 3D tissues
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Citation
Miller, Jordan S., Kelly R. Stevens, Michael T. Yang, Brendon M.
Baker, Duc-Huy T. Nguyen, Daniel M. Cohen, Esteban Toro, et
al. “Rapid Casting of Patterned Vascular Networks for Perfusable
Engineered Three-Dimensional Tissues.” Nature Materials 11,
no. 9 (July 1, 2012): 768–774.
As Published
http://dx.doi.org/10.1038/nmat3357
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Nature Publishing Group
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Author's final manuscript
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Thu May 26 01:57:38 EDT 2016
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http://hdl.handle.net/1721.1/85973
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Published in final edited form as:
Nat Mater. 2012 September ; 11(9): 768–774. doi:10.1038/nmat3357.
Rapid casting of patterned vascular networks for perfusable
engineered 3D tissues
Jordan S. Millera, Kelly R. Stevensb, Michael T. Yanga, Brendon M. Bakera, Duc-Huy T.
Nguyena, Daniel M. Cohena, Esteban Toroa, Alice A. Chenb, Peter A. Galiea, Xiang Yua,
Ritika Chaturvedia, Sangeeta N. Bhatiab,c,d, and Christopher S. Chena,e
aDepartment of Bioengineering, University of Pennsylvania, Philadelphia, PA 19104
bHarvard-MIT
Division of Health Sciences and Technology, Massachusetts Institute of
Technology, Cambridge, MA 02139
cHoward
Hughes Medical Institute, Cambridge, MA 02139
dDepartment
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of Electrical Engineering and Computer Science, Massachusetts Institute of
Technology, Cambridge, MA 02139
Abstract
In the absence of perfusable vascular networks, three-dimensional (3D) engineered tissues densely
populated with cells quickly develop a necrotic core [1]. Yet the lack of a general approach to
rapidly construct such networks remains a major challenge for 3D tissue culture [2–4]. Here, we
3D printed rigid filament networks of carbohydrate glass, and used them as a cytocompatible
sacrificial template in engineered tissues containing living cells to generate cylindrical networks
which could be lined with endothelial cells and perfused with blood under high-pressure pulsatile
flow. Because this simple vascular casting approach allows independent control of network
geometry, endothelialization, and extravascular tissue, it is compatible with a wide variety of cell
types, synthetic and natural extracellular matrices (ECMs), and crosslinking strategies. We also
demonstrated that the perfused vascular channels sustained the metabolic function of primary rat
hepatocytes in engineered tissue constructs that otherwise exhibited suppressed function in their
core.
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Living tissues have complex mass transport requirements that are principally met by blood
flow through multiscale vascular networks of the cardiovascular system. Such vessels
deliver nutrients and oxygen to, and remove metabolic byproducts from, all of the organ
systems in the body and were critical to the rise of large-scale multicellular organisms [5].
Although tremendous progress has been made in the past several decades to isolate and
culture cells from native tissues, simple methods to generate tissue constructs populated at
physiologic cell densities that are sustained by even the most basic vascular architectures
have remained elusive.
To create perfusable channels in engineered tissues, layer-by-layer assembly [6–9] has been
explored. In this approach, a trench is molded into one layer such that a second, separately
fabricated layer can then be aligned and laminated to close the lid to form channels in an
iterative fashion. However, layer-by-layer assembly is slow and results in seams or other
e
To whom correspondence should be addressed. chrischen@seas.upenn.edu.
Author Contributions
J.S.M. and C.S.C. conceived and initiated the project. J.S.M., K.R.S., M.T.Y., B.M.B., D-H.T.N., D.M.C., E.T., A.A.C., P.A.G., X.Y.,
and R.C. designed and performed experiments. C.S.C. and S.N.B. supervised the project.
Miller et al.
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structural artifacts throughout the construct while simultaneously placing considerable
design constraints on the materials, channels, and cells used during fabrication. Bioprinting
[10], in which cells and matrix are deposited dropwise, has been developed over the past
decade but also is a slow, serial process with limitations on print resolution, materials, and
cells. In contrast to these methods, 3D sacrificial molding [11–13] provides an intriguing
alternative. Proof-of-concept studies have shown that a network of channels can be
fabricated by creating a rigid 3D lattice of filaments, casting the lattice into a rubber or
plastic material, and then sacrificing the lattice to reveal a microfluidic architecture in the
bulk material. However, 3D sacrificial molding of perfusable channels has so far required
the use of cytotoxic organic solvents or processing conditions for either removing the
sacrificial filaments or casting the surrounding material, and thus could not be accomplished
with aqueous-based ECMs or in the presence of living cells.
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Here, we describe a biocompatible sacrificial material—a simple glass made from mixtures
of inexpensive and readily available carbohydrates—and a means to 3D print the material to
facilitate the rapid casting of patterned vascular networks in engineered tissues. This
carbohydrate glass formulation was developed specifically to accommodate two seemingly
opposing design criteria that we identified for biocompatible 3D sacrificial materials: 1)
sufficient mechanical stiffness to physically support its own weight in an open 3D lattice of
filaments and 2) the ability to dissolve rapidly and biocompatibly in the presence of living
cells.
Carbohydrate glass can be formed by dissolving one or more carbohydrates in water and
then boiling of the solvent. Our early experiments were based on a sucrose-glucose mixture
developed by the food industry, which showed that while sucrose is unstable in
supersaturated solutions, the addition of glucose prevents recrystallization and facilitates the
formation of a stable and inexpensive glass [14]. This simple mixture was too hygroscopic
and soft to handle. During material optimization and screening of potential additives, we
found that the addition of starch stiffened the base material, but it imparted inferior optical
clarity and therefore limited potential use with matrices that are commonly crosslinked by
photochemical reactions [15, 16]. In contrast, the addition of glycerol preserved clarity but
rendered filaments mechanically unstable at room temperature. Ultimately, we further
reinforced the glass and improved its temperature stability by incorporating dextrans.
Uniaxial compression testing confirmed that the carbohydrate glass is mechanically stiff and
brittle at room temperature (Figure 1a). The optical transparency of the glass indicated
compatibility with photopolymerization wavelengths in the ultraviolet, visible, and nearinfrared ranges, making it unlikely that the glass would leave shadowing artifacts in
photoactive scaffolds (Figure 1b).
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Multiscale vascular networks are comprised of a range of diameters of vessels and their
interconnections. Thermal extrusion and fiber drawing with a 3D printer—a programmable
Cartesian coordinate positioning system—provided an effective route to the fabrication of
filamentous carbohydrate glass lattices. By varying only the translational velocity of the
extrusion nozzle, while holding constant the nozzle diameter and the extrusion flow rate
parameters, extruded filament diameters tracked the governing equation:
(1)
where D(v) is the resultant filament diameter, A is a constant that incorporates the extrusion
nozzle diameter and extrusion flow rate, and v is the velocity of the extrusion nozzle (Figure
1c). This relationship derives from existing models of glass fiber drawing [17] and allowed
the generation of carbohydrate glass lattices in predefined, multiscale, and reproducible
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patterns (Figure 1d–h). Moreover, controlling the temperature of the assembly platform
facilitated the formation of smooth melt fusions at filament intersections (Figure 1e).
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We next sought to use these lattices as a sacrificial element for creating fluidic channels
within monolithic cellularized tissue constructs (Figure 2a). In our strategy, a suspension of
cells in ECM prepolymer is poured to encapsulate the lattice. After crosslinking the ECM,
the glass filaments are dissolved to form vessels while their interfilament fusions become
intervessel junctions (Figure 2b–c). To prevent disruption of ECM crosslinking and to avoid
the potential for osmotic damage to encapsulated cells due to carbohydrate dissolution, we
coated the carbohydrate glass lattice with a thin layer of poly(D-lactide-co-glycolide)
(PDLGA) prior to casting ECM. This coating allowed the dissolved carbohydrates to be
owed out of the formed channels instead of through the bulk of the engineered construct
(Figure 2b and Supplementary Movie 1). Importantly, the coating did not inhibit the ability
of the network to support convective and diffusive transport into the bulk gel (Figure 2c).
Additionally, we observed that, after sacrifice, the glass interfilament fusions left behind
smooth elliptical intervessel junctions that supported fluidic connection between adjoining
vascular channels.
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To demonstrate the flexibility and generality of this approach, we patterned vascular
channels in the presence of living cells in a wide range of natural and synthetic ECM
materials (Figure 2d). The time required for encapsulating cells and lattices in ECM
prepolymer, ECM crosslinking, and glass dissolution is on the order of minutes.
Importantly, we chose ECM materials which varied not only in their bulk material properties
but also in their means of crosslinking. Indeed, the approach generated channels without the
need to modify handling of aqueous cellularized gels formed by chain entanglements
(cooling of agarose), ionic interactions (calcium-polymerized alginate), photopolymerization
(synthetic PEG-based hydrogels, [18]), enzymatic activity (thrombin-polymerized fibrin),
and protein precipitation (warming of Matrigel). As predicted from characterization of the
optical transparency of the carbohydrate glass, photopolymerized gels exhibited no visible
shadowing artifacts due to light absorption by the patterned glass lattice. To our knowledge,
no other channel-forming technique is compatible with such a wide range of ECM materials.
The approach also appears to have no negative effects on cells. Encapsulated cells survived,
spread, and migrated in channeled scaffolds at levels not different from non-channeled
control gels, demonstrating biocompatibility of the entire vessel casting process (Figure 2e).
Similar viability was found for human umbilical vein endothelial cells (HUVECs), 10T1/2
cells, human fibroblasts, and human embryonic kidney (HEK) cells (data not shown).
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Due to the mechanical rigidity and self-supporting nature of the carbohydrate glass lattice,
introducing 3D multilayer architecture into the engineered vasculature requires no additional
constraints, time, or steps to the sacrificial process (Supplementary Figure S1a). To
demonstrate the compatibility of this vascular casting approach with additional design
considerations often important to the engineering of tissues, we fabricated complex cellular
and immobilized factor gradients in tissue constructs that also contained our channels
(Supplementary Figure S1b). Analysis of these tissue constructs (Supplementary Figure S1c)
demonstrated that patterning of cells or immobilized factors within the construct into step,
linear, and exponential gradients could be accomplished in a single engineered tissue
construct containing perfusable vascular channels. Here, cells were encapsulated along with
immobilized fluorescent beads as a model factor but this technology should be readily
translated to immobilized gradients of adhesive peptides [19], proteins and growth factors
[20–22], or the ECM itself [23, 24]. Together, these results illustrate that the available
design parameter space for a tissue construct is, for the first time, unhindered by the
inclusion of patterned vascular channels and junctions.
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Vascularized solid tissues can be conceptually reduced to a “vascular unit cell” comprised of
three key compartments: 1) the vascular lumen which serves as both the source and sink for
most soluble factors, 2) endothelial cells lining the vascular wall which regulate mass
transport exchange with the interstitium, and 3) cells and matrix residing in the interstitial
zone between vascular channels (Figure 3a). Here, we demonstrate control over each of
these compartments in engineered tissue constructs. The monolithic nature of the gels
(resulting from a single step polymerization) and the lack of architectural seams supported
non-leaking perfusion of human blood under positive pressure with either laminar or
turbulent pulsatile flow (Figure 3b and Supplementary Movies 2 and 3), with smooth
interchannel junctions supporting branched fluid flow. Endothelial cells seeded through a
single inlet in the network quickly lined the walls of the entire network, including the
junctions between vessels of differing diameters (Figure 3c,d). Because this endothelium is
formed after forming the tissue, these cells and their seeding are independently introduced
from cells encapsulated in the interstitial zone. In co-cultures with 10T1/2 cells in the
interstitial space, endothelial cells lining the vascular lumen became surrounded by the
10T1/2 cells and formed single and multicellular sprouts extending from the patterned
vasculature into the bulk gel (Figure 3e,f).
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Engineered constructs densely populated with cells can develop a necrotic core due to lack
of adequate mass transport [1]. Thus, a major functional requirement of such an engineered
vasculature is its utility to sustain cellular activity in metabolically demanding settings such
as physiologically high cell densities. To facilitate imaging of such a densely populated
construct, we generated a single layer of parallel fluidic channels formed within the center of
a rectangular construct. As a measure of cellular function and activity, we examined
expression of destabilized EGFP from a constitutively expressed lentiviral cassette inserted
into HEK293T cells. Monolithic slab gels uniformly distributed with cells exhibited cellular
activity only at the gel perimeter (Figure 4a). In contrast, gels with channels rescued protein
expression in the gel core, most dramatically around each perfused channel. This
preservation of cell function was perfusion-dependent, as EGFP expression was absent near
channels without perfusion (data not shown). We also looked at a broader range of cellular
densities in these constructs with a functional enzyme assay of a constitutively secreted
Gaussia luciferase reporter. At low cell densities, diffusion alone is able to maintain cellular
function in bulk gels (Figure 4b). As nutrient requirements of the tissue construct increased
with cell density, Gaussia production began to plateau. This limitation was overcome by
convective transport through the channeled scaffold. The maintenance of cellular metabolic
activity (Figure 4a) as well as secretion of functional proteins and enzymes (Figure 4b) are
important functional outputs for many types of engineered tissues [25], both of which appear
to be maintained with our engineered vasculature. Together these results illustrate the
capacity of patterned perfusable channel architectures to provide functional mass transport
to 3D cell cultures at or near physiologic cell densities.
Although these data demonstrate the utility of the approach in supporting a transformed cell
line, primary parenchymal cells that would ultimately be required for clinically implanted
engineered tissues often cannot tolerate stresses associated with extended periods of
suspension and hypoxia. We therefore engineered perfusable gels containing primary
hepatocytes, which are known to be highly sensitive to hypoxia and handling (Figure 4c–d).
After 8 days in culture, perfusable tissues exhibited substantially higher albumin secretion
and urea synthesis compared to slab gels (gels without channels) of the same volume (Figure
4d). Optical sections of these constructs showed that at high cell concentrations cell survival
was most prevalent adjacent to perfused channels and decayed radially, consistent with
patterns observed in the dsEGFP-expressing HEK293T cells as well as a 3D finite-element
model of nutrient delivery to the entrapped cells (Supplementary Figure S2). Together these
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results illustrate the utility of this strategy for supporting the function of engineered tissues
comprised of even highly sensitive primary cells.
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Existing methods to create cell-laden gels containing a microfluidic network have required
the delicate process of precise stacking and lamination of individually fabricated layers [6–
9]. In this study, we found sacrificial carbohydrate glass lattices to be well-suited for the
creation of densely populated tissue constructs with perfusable vascular channels and
junctions. A key advantage of our method is that the entire perfusable scaffold is formed as a
continuous phase simply by filling the 3D void volume around carbohydrate glass lattices
and crosslinking the matrix. Cells are encapsulated in ECM and the resulting tissue construct
can be perfused within minutes. We believe the sheer rapidity of this process prevents the
formation of a necrotic core for metabolically demanding cells during fabrication. Moreover,
the microstructural characteristics of the fluidic network—such as vessel diameter,
circularity, surface roughness, and junction architecture—arise from fiber drawing and
surface tension rather than photolithography or micromachining. Relying on simple physical
principles, rather than on engineering processes, enabled the use of low precision hardware
to rapidly and reproducibly generate multiscale microvascular architectures in aqueousbased biomaterial scaffolds containing living cells.
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Further, the process separation between the 3D microfabrication of filament networks and
the handling of cells and ECM allows for dissemination of the technology to distant research
laboratories. To illustrate this feature in the current study, carbohydrate glass lattices were
shipped under ambient conditions between laboratories. Primary liver hepatocytes were then
encapsulated via standard manual pipetting steps to rapidly create perfusable hepatic tissues
or non-perfusable control gels. The ability to access this vascularization strategy without
fabricating the networks in-house may facilitate rapid adoption of the technology.
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Engineered 3D constructs have gained increased attention as in vitro tools for the study of
cell-cell and cell-matrix interactions, and are being explored for potential use as
experimental models or therapeutic replacements of human tissues. However, with the
exception of avascular or thin tissues, it has been difficult to achieve the cellular densities of
native tissues (approximately 10–500 million cells per milliliter). The approach described
here demonstrates an avenue for building and studying such tissue mimics, in which the
vasculature appears not to constrain the design space for the tissue itself, allowing for
arbitrary cell types, matrices, and their patterning. Coupled with advances in microfluidic
device technologies [26, 27], the controlled architecture of these engineered fluidic networks
may also provide a means to directly examine the interplay between mass transport
requirements of specific tissues and vascular architecture. Moreover, the facile and highly
automated nature of this perfusable tissue fabrication strategy should provide a flexible
platform for a wide array of specific applications, and may enable the scaling of densely
populated tissue constructs to arbitrary size.
1. Methods
1.1. Preparation and 3D printing of carbohydrate glass
A mixture of 25 g glucose, 53 g sucrose, 10 g dextran (86 kDa), and 50 mL reverse osmosis
water (≥18 MΩ; Millipore) was warmed to 165 °C to remove most of the water and form a
liquid glass. The hot mixture was poured into a 50 mL syringe that was maintained at 110
°C. The syringe was mounted on a custom modified RepRap Mendel 3D printer with
associated electronics (Gen3, MakerBot; RAMPS + RAMMS, Ultimachine). Custom Python
scripts were developed to generate the 3D motion control GCode used to drive the machine
via open-source ReplicatorG software. Carbohydrate glass lattices were 3D printed at 110
°C under nitrogen pressure with pneumatic control through a 16-gauge or 18-gauge (1.2 mm
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or 0.84 mm ID, respectively) steel nozzle, vitrified to 50 °C, and then immersed in a 25 mg/
mL solution of poly(D-lactide-co-glycolide) (PDLGA; Purac, Amsterdam) in chloroform for
up to 5 min. Glass lattices were encapsulated in ECM along with living cells on the same
day the lattices were fabricated, or were stored at 45 °C or in a vacuum chamber until use to
protect the hygroscopic carbohydrates from absorbing ambient moisture under atmospheric
conditions.
1.2. Polymer synthesis and cell and lattice encapsulation
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Poly(ethylene glycol) diacrylate (PEGDA, 6 kDa or 35 kDa) and acrylate-PEG-RGDS (4
kDa) were synthesized as previously described [18]. In a typical experiment, a prepolymer
mixture containing PEGDA (5, 10, or 20 wt%), acrylate-PEG-RGDS (1 mM), photoinitiator
(Irgacure 2959, Ciba Geigy, 0.05% w/v), and cells of interest (1–40e6 cells/mL) was
dispensed into rectangular molds containing suspended carbohydrate glass lattices (500 µL
total volume per gel). PEG hydrogels were photopolymerized (Omnicure 2000, 320–500
nm) at 100 mW/cm2 for two repetitions of 30 sec duration (rotating 180° about the y-axis
before the second exposure). Fibrin gels (10–40 mg/mL) were created by combining
fibrinogen, thrombin and cell suspension in phosphate buffered saline (PBS) and then
dispensing this mixture around a carbohydrate glass lattice. Fibrin gels were polymerized for
10 min at 37 °C or for 20 min at room temperature. Matrigel constructs were formed by
mixing a cell suspension with Matrigel and then dispensing the mixture around a
carbohydrate glass lattice, followed by incubation at 37 °C for 10 min. Alginate gels (2%)
were formed by mixing an alginate solution with cell suspension, dispensing the mixture
around a carbohydrate glass lattice, then carefully crosslinked with a 50 mg/mL CaCl2
solution for 10 min. Agarose gels (2%) were formed by mixing a low-melt agarose solution
with cell suspension, dispensing the mixture around a carbohydrate glass lattice, then placed
at 4 °C for 20 min. All crosslinked gels were post-processed identically: after crosslinking
they were placed in complete medium to dissolve carbohydrate glass (10 min), followed by
exchange with fresh medium and cell culture (static culture or orbital shaking at a rate of 2
Hz). In 800 µm channels (diameter) we measured peak flow rates of 10 µL/sec,
corresponding to mean velocities of 5 mm/sec and shear stress of 1 dyn/cm2. These values
are comparable to physiologic settings. Gel slabs (without vascular architectures) were
created by using identical rectangular molds and identical gel processing steps, but without
encapsulating glass lattices. Diffusion studies were conducted with fluorescent Cascade Blue
Dextran 10kDa (Invitrogen). Fluorescent beads (Polysciences) were mixed with ECM
prepolymer mixtures for matrix immobilization or PBS for bead perfusion studies.
Heparanized human blood, whole blood, or packed red blood cells (Interstate Blood Bank)
were washed and diluted with PBS before use.
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1.3. Endothelialization of vascular networks
HUVECs were seeded in the vascular lumen by injecting a HUVEC suspension (35e6 cells/
mL) into the vascular architecture approximately 10 minutes after the constructs were
fabricated. HUVECs were allowed to attach in static culture for one hour before introducing
flow, and they reached confluence within 1 day. Endothelialized gels often contained
additional HUVECs (1.5e6/mL) and 10T1/2 cells (1.5e6/mL) encapsulated in the bulk gel,
such as the gel shown in Figure 3c–f.
Online Methods
Additional methods may be found online in Supplementary Information.
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Supplementary Material
Refer to Web version on PubMed Central for supplementary material.
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Acknowledgments
We thank the large number of open source and related projects that critically facilitated this work, including
Arduino.cc, RepRap.org, MakerBot.com, Replicat.org, MakerGear.com, Ultimachine.com, Hive76.org, Python.org,
Hugin.SourceForge.net, ImageMagick.org, Blender.org, Enblend.sourceforge.net, NIH ImageJ, and Fiji.sc. We
thank Robert J. Vlacich and Christopher D. Thompson for assistance with precision pneumatic extrusion, Aaron
Dominguez for assistance with red blood cell isolation, and Yi-Ju Chen for assistance with transduction. This work
was supported in part by grants from the US National Institutes of Health (EB00262, EB08396, GM74048), the
Penn Center for Engineering Cells and Regeneration, and the American Heart Association-Jon Holden DeHaan
Foundation. Individual fellowship support was provided by Ruth L. Kirschstein National Research Service Awards
from NIH (J.S.M., HL099031; K.R.S., DK091007), the National Science Foundation IGERT program (M.T.Y.,
DGE-0221664), and the American Heart Association (X.Y., 10POST4220014).
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Figure 1.
Carbohydrate glass material properties and filament architecture formation. (a) Stress-strain
curve from uniaxial compression testing indicates the carbohydrate glass is a stiff and brittle
material at 25 °C, with Young’s modulus = 1 GPa (measured in the linear regime),
maximum strength of 28 MPa, maximum strain of 3.25%. (b) Optical extinction for a 1 cm
sample of carbohydrate glass indicates the material transmits light wavelengths commonly
used during biocompatible imaging and photopolymerization (365–550 nm, shaded box). (c)
During thermal extrusion and 3D printing, filament diameter is controlled by the travel
speed of the extrusion nozzle and follows a simple power law from glass fiber drawing
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(equation inset). (d) Architectural design of a multiscale carbohydrate glass lattice (green).
(e) Top view of the multiscale architectural design in (d) 3D printed in carbohydrate glass
(scale bar = 1 mm). Interfilament melt fusions are magnified and shown in side-view (scale
bars = 200 µm). (f) Multilayered lattices are fabricated in minutes with precise lateral and
axial positioning resolution (scale bar = 1 mm). (g) A multiscale architecture showing a
single 1 mm filament (top) connected to angled arrays of smaller interconnected filaments
(scale bar = 1 mm). (h) Serial y-junctions and curved filaments can also be fabricated (scale
bars = 1 mm).
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Figure 2.
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Monolithic tissue construct containing patterned vascular architectures and living cells. (a)
Schematic overview. An open, interconnected, self-supporting carbohydrate glass lattice is
3D printed to serve as the sacrificial element for the casting of vascular architectures. The
lattice is encapsulated in ECM along with living cells. The lattice is dissolved in minutes in
cell media without damage to nearby cells. The process yields a monolithic tissue construct
with a vascular architecture that matches the original lattice. (b) A single carbohydrate glass
fiber (200 µm in diameter, top) is encapsulated in a fibrin gel. Following ECM crosslinking,
the gel and filament are immersed in aqueous solution and the dissolved carbohydrates are
flowed out of the resulting channel (middle). Removal of the filament yields an open
perfusable channel in the fibrin gel (bottom, scale bar = 500 µm). See Supplementary
Movie 1 for full time course. (c) A fibrin gel with patterned interconnected channels of
different diameters supports convective and diffusive transport of a fluorescent dextran
injected into the channel network (upper left, phase contrast, scale bar = 500 µm). Line plot
of normalized fluorescence across the gel and channel (blue arrow) shows a sinusoidal
profile in the channel (between dotted black lines) characteristic of a cylinder and temporal
diffusion from the channel into the bulk gel. Enlargement of the dotted box region shows an
oval intervessel junction between the two perpendicular channels (right, scale bar = 100
µm). (d) Cells constitutively expressing EGFP were encapsulated (5e6/mL) in a variety of
ECM materials then imaged with confocal microscopy to visualize the matrix (red beads),
cells (10T1/2, green) and the perfusable vascular lumen (blue beads) shown schematically
(bottom right). The materials have varied crosslinking mechanisms (annotated above
images) but were all able to be patterned with vascular channels. Scale bars = 200 µm. (e)
Representative cross-section image of unlabeled HUVEC (1e6/mL) and 10T1/2 (1e6/mL)
co-cultures (not expressing EGFP) encapsulated uniformly in the interstitial space of fibrin
gel (10 mg/mL) with perfusable networks after two days in culture were stained with a
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fluorescent Live/Dead assay (green, Calcein AM; red, Ethidium Homodimer). Cells survive
and spread near open cylindrical channels (highlighted with white arrow). Scale bar = 200
µm.
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Figure 3.
Demonstrated control over the three key compartments of vascularized solid tissues, shown
schematically in (a) as the “vascular unit cell” comprised of the vascular lumen, endothelial
cells lining the vascular wall, and the interstitial zone containing matrix and encapsulated
cells. (b) Patterned vascular channels support positive pressure and pulsatile flow of human
blood with intervessel junctions supporting branched fluid flow (left). Spiral flow patterns
(right, 0.4 sec) are characteristic of non-laminar flow through cylindrical channels. See
Supplementary Movies 2 and 3, Scale bars = 1 mm, left; 2 mm, right. (c) Control of the
interstitial zone and the lining endothelium of vascularized tissue constructs is demonstrated
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by encapsulating 10T1/2 cells (1.5e6/mL, constitutively expressing EGFP) in the interstitial
space of a fibrin gel (10 mg/mL) followed by seeding of HUVECs (constitutively expressing
mCherry) throughout the vascular network via a single lumenal injection (see Methods).
After one day in culture a confocal z-stack montage demonstrated HUVECs residing in the
vascular space with 10T1/2 uniformly distributed throughout the bulk gel. Scale bar = 1 mm.
(d) A partial z-stack of two intersecting channels demonstrated endothelialization of channel
walls and across the intervessel junction, while in the surrounding bulk gel 10T1/2 cells are
seen beginning to spread out in 3D. See Supplementary Movie 4 for the complete 700 µm
z-stack from (d). (e) After 9-days in culture, cross-section imaging of a representative
channel (optical thickness and z-position = 10 µm) demonstrated that the endothelial
monolayer lining the vascular lumen became surrounded by 10T1/2 cells. Scale bar = 200
µm. (f) Endothelial cells formed single and multicellular sprouts (arrowheads) from
patterned vasculature as seen in a z-stack (optical thickness = 200 µm) from deeper within
the gel (z-position = 300 µm, left). Even deeper imaging (z-position = 950 µm, optical
thickness = 100 µm, right) confirmed that the vascular lumen remained open throughout
vessels and intervessel junctions and that endothelial cells also sprouted from larger vessels
(arrowheads). See Supplementary Movie 5 for the complete 1 mm z-stack from (e,f).
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Figure 4.
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Perfusion channels sustain cellular metabolic function in the core of thick, densely populated
tissue constructs. (a) Representative cross-section image montages of PEG hydrogels
containing 40e6 HEK cells/mL after three days in culture. The intracellular dsEGFP reporter
spatially indicates cells are active at the gel slab perimeter and circumferentially around
perfusion channels, but not elsewhere in the gel core. Scale bar = 2 mm. (b) A functional
enzyme assay of secreted Gaussia luciferase from these constructs indicates that the channel
architecture preserves cell function even at high cell densities, where function in slab gels
quickly falls off. (c) Primary rat hepatocytes (24e6 hepatocytes/mL) and stabilizing stromal
fibroblasts in agarose gels (slab vs. channeled) after 8 days of culture were stained with a
fluorescent Live/Dead assay (green, Calcein AM; red, Ethidium Homodimer). Cells survive
at the gel perimeter and near perfused channels, and survival decayed deeper in the gels.
Scale bar = 1 mm. (d) Assessment of albumin secretion (top) and urea synthesis (bottom)
by primary hepatocytes (16e6/mL) in gels after 8 days of culture demonstrated improved
hepatic function in channeled gels compared to slab gels. Error bars represent standard error,
p-values for channel versus slab gel comparisons: * < 0.0051; ‡ < 0.045.
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